CRISPR-Cas9 engineering of the RAG2 locus via complete coding sequence replacement for therapeutic applications.

RAG2-SCID is a primary immunodeficiency caused by mutations in Recombination-activating gene 2 (RAG2), a gene intimately involved in the process of lymphocyte maturation and function. ex-vivo manipulation of a patient's own hematopoietic stem and progenitor cells (HSPCs) using CRISPR-Cas9/rAAV6 gene editing could provide a therapeutic alternative to the only current treatment, allogeneic hematopoietic stem cell transplantation (HSCT). Here we show an innovative RAG2 correction strategy that replaces the entire endogenous coding sequence (CDS) for the purpose of preserving the critical endogenous spatiotemporal gene regulation and locus architecture. Expression of the corrective transgene leads to successful development into CD3+TCRαß+ and CD3+TCRγδ+ T cells and promotes the establishment of highly diverse TRB and TRG repertoires in an in-vitro T-cell differentiation platform. Thus, our proof-of-concept study holds promise for safer gene therapy techniques of tightly regulated genes.

Multiplex HDR for disease and correction modeling of SCID by CRISPR genome editing in human HSPCs.

Severe combined immunodeficiency (SCID) is a group of disorders caused by mutations in genes involved in the process of lymphocyte maturation and function. CRISPR-Cas9 gene editing of the patient's own hematopoietic stem and progenitor cells (HSPCs) ex vivo could provide a therapeutic alternative to allogeneic hematopoietic stem cell transplantation, the current gold standard for treatment of SCID. To eliminate the need for scarce patient samples, we engineered genotypes in healthy donor (HD)-derived CD34+ HSPCs using CRISPR-Cas9/rAAV6 gene-editing, to model both SCID and the therapeutic outcomes of gene-editing therapies for SCID via multiplexed homology-directed repair (HDR). First, we developed a SCID disease model via biallelic knockout of genes critical to the development of lymphocytes; and second, we established a knockin/knockout strategy to develop a proof-of-concept single-allelic gene correction. Based on these results, we performed gene correction of RAG2-SCID patient-derived CD34+ HSPCs that successfully developed into CD3+ T cells with diverse TCR repertoires in an in vitro T cell differentiation platform. In summary, we present a strategy to determine the optimal configuration for CRISPR-Cas9 gene correction of SCID using HD-derived CD34+ HSPCs, and the feasibility of translating this gene correction approach in patient-derived CD34+ HSPCs.

Different hotspot p53 mutants exert distinct phenotypes and predict outcome of colorectal cancer patients.

The TP53 gene is mutated in approximately 60% of all colorectal cancer (CRC) cases. Over 20% of all TP53-mutated CRC tumors carry missense mutations at position R175 or R273. Here we report that CRC tumors harboring R273 mutations are more prone to progress to metastatic disease, with decreased survival, than those with R175 mutations. We identify a distinct transcriptional signature orchestrated by p53R273H, implicating activation of oncogenic signaling pathways and predicting worse outcome. These features are shared also with the hotspot mutants p53R248Q and p53R248W. p53R273H selectively promotes rapid CRC cell spreading, migration, invasion and metastasis. The transcriptional output of p53R273H is associated with preferential binding to regulatory elements of R273 signature genes. Thus, different TP53 missense mutations contribute differently to cancer progression. Elucidation of the differential impact of distinct TP53 mutations on disease features may make TP53 mutational information more actionable, holding potential for better precision-based medicine.

High-Throughput Imaging of CRISPR- and Recombinant Adeno-Associated Virus-Induced DNA Damage Response in Human Hematopoietic Stem and Progenitor Cells.

CRISPR-Cas technology has revolutionized gene editing, but concerns remain due to its propensity for off-target interactions. This, combined with genotoxicity related to both CRISPR-Cas9-induced double-strand breaks and transgene delivery, poses a significant liability for clinical genome-editing applications. Current best practice is to optimize genome-editing parameters in preclinical studies. However, quantitative tools that measure off-target interactions and genotoxicity are costly and time-consuming, limiting the practicality of screening large numbers of potential genome-editing reagents and conditions. Here, we show that flow-based imaging facilitates DNA damage characterization of hundreds of human hematopoietic stem and progenitor cells per minute after treatment with CRISPR-Cas9 and recombinant adeno-associated virus serotype 6. With our web-based platform that leverages deep learning for image analysis, we find that greater DNA damage response is observed for guide RNAs with higher genome-editing activity, differentiating even single on-target guide RNAs with different levels of off-target interactions. This work simplifies the characterization and screening process of genome-editing parameters toward enabling safer and more effective gene-therapy applications.

Microfluidic tool for rapid functional characterization of CRISPR complexes.

RNA guided nucleases are regarded as the future genome editing technologies. As such, they need to meet strong safety margins. Two major challenges in incorporating CRISPR technologies into the clinical world are off-target activity and editing efficiency. The common way to tackle such issues is to measure the binding and cleavage kinetics of the CRISPR enzyme. This can be challenging since, for example, DNA is not released from the CAS9 protein post cleavage. Here a promising new microfluidic approach to characterizing Enzymatic Interaction and Function of CRISPR complexes on a microfluidic platform (EnzyMIF) is presented. The method can rapidly detect the kd, koff, km and kcat for various RNA guided nucleases. In this work, two single guide RNAs with significantly different in-cell cleavage efficiency, RAG2 and RAG1, are used as proof-of-concept. The EnzyMIF assay results provide biochemical characterization of these guide RNAs that can explain the difference in cleavage using both wild type (WT) CAS9 and HiFi CAS9. Notably, it is shown that EnzyMIF characterization correlates with cell culture genomic editing efficiency results. It is suggested that EnzyMIF can predict the quality of cleavage rapidly and quantitatively.

CRISPECTOR provides accurate estimation of genome editing translocation and off-target activity from comparative NGS data.

Controlling off-target editing activity is one of the central challenges in making CRISPR technology accurate and applicable in medical practice. Current algorithms for analyzing off-target activity do not provide statistical quantification, are not sufficiently sensitive in separating signal from noise in experiments with low editing rates, and do not address the detection of translocations. Here we present CRISPECTOR, a software tool that supports the detection and quantification of on- and off-target genome-editing activity from NGS data using paired treatment/control CRISPR experiments. In particular, CRISPECTOR facilitates the statistical analysis of NGS data from multiplex-PCR comparative experiments to detect and quantify adverse translocation events. We validate the observed results and show independent evidence of the occurrence of translocations in human cell lines, after genome editing. Our methodology is based on a statistical model comparison approach leading to better false-negative rates in sites with weak yet significant off-target activity.

Chemical Modification of Guide RNAs for Improved CRISPR Activity in CD34+ Human Hematopoietic Stem and Progenitor Cells.

Human CD34+ hematopoietic stem and progenitor cells (HSPCs) have the unique ability to repopulate the entire hematopoietic system and thus are at the center of diverse, therapeutically relevant studies. The recent development of the CRISPR/Cas9 tool made the powerful research technique of genome editing highly accessible. Our previous studies demonstrated that high editing efficiency is reached when the CRISPR/Cas9 is introduced to CD34+ HSPCs as a ribonucleoprotein (RNP) complex with chemically modified guide RNAs (gRNAs). The current protocol details a quick 4-day procedure for ex vivo genome editing in CD34+ HSPCs by RNP complexes that are targeted to a specific locus by either a single gRNA (sgRNA) or a 2-part gRNA. The delivery of RNP complexes is performed by electroporation in the presence of a nonspecific, ssDNA electroporation enhancer, which highly improves editing efficiency under the described conditions. This approach is simple and effective with the potential to accelerate many biotechnological and therapeutic applications of the CRISPR/Cas9 technology.

Increasing CRISPR Efficiency and Measuring Its Specificity in HSPCs Using a Clinically Relevant System.

Genome editing of human cluster of differentiation 34+ (CD34+) hematopoietic stem and progenitor cells (HSPCs) holds great therapeutic potential. This study aimed to optimize on-target, ex vivo genome editing using the CRISPR-Cas9 system in CD34+ HSPCs and to create a clear workflow for precise identification of off-target effects. Modified synthetic guide RNAs (gRNAs), either 2-part gRNA or single-guide RNA (sgRNA), were delivered to CD34+ HSPCs as part of ribonucleoprotein (RNP) complexes, targeting therapeutically relevant genes. The addition of an Alt-R electroporation enhancer (EE), a short, single-stranded oligodeoxynucleotide (ssODN), significantly increased editing efficiency in CD34+ HSPCs. Notably, similar editing improvement was observed when excess gRNA over Cas9 protein was used, providing a DNA-free alternative suitable for therapeutic applications. Furthermore, we demonstrated that sgRNA may be preferable over 2-part gRNA in a locus-specific manner. Finally, we present a clear experimental framework suitable for the unbiased identification of bona fide off-target sites by Genome-Wide, Unbiased Identification of Double-Strand Breaks (DSBs) Enabled by Sequencing (GUIDE-seq), as well as subsequent editing quantification in CD34+ HSPCs using rhAmpSeq. These findings may facilitate the implementation of genome editing in CD34+ HSPCs for research and therapy and can be adapted for other hematopoietic cells.

WWOX and p53 Dysregulation Synergize to Drive the Development of Osteosarcoma.

Osteosarcoma is a highly metastatic form of bone cancer in adolescents and young adults that is resistant to existing treatments. Development of an effective therapy has been hindered by very limited understanding of the mechanisms of osteosarcomagenesis. Here, we used genetically engineered mice to investigate the effects of deleting the tumor suppressor Wwox selectively in either osteoblast progenitors or mature osteoblasts. Mice with conditional deletion of Wwox in preosteoblasts (WwoxΔosx1) displayed a severe inhibition of osteogenesis accompanied by p53 upregulation, effects that were not observed in mice lacking Wwox in mature osteoblasts. Deletion of p53 in WwoxΔosx1 mice rescued the osteogenic defect. In addition, the Wwox;p53Δosx1 double knockout mice developed poorly differentiated osteosarcomas that resemble human osteosarcoma in histology, location, metastatic behavior, and gene expression. Strikingly, the development of osteosarcomas in these mice was greatly accelerated compared with mice lacking p53 only. In contrast, combined WWOX and p53 inactivation in mature osteoblasts did not accelerate osteosarcomagenesis compared with p53 inactivation alone. These findings provide evidence that a WWOX-p53 network regulates normal bone formation and that disruption of this network in osteoprogenitors results in accelerated osteosarcoma. The Wwox;p53Δosx1 double knockout establishes a new osteosarcoma model with significant advancement over existing models. Cancer Res; 76(20); 6107-17. ©2016 AACR.